Purpose:
To investigate the possible antiapoptotic effect of acetylcholine (ACh) in Fas-mediated apoptosis of primary human keratocytes in vitro, and to explore the underlying mechanism.

Methods:
Primary human keratocytes were isolated from healthy corneas. Fas ligand (FasL) was used to induce apoptosis in keratocytes. Cell death was assessed by ELISA. Activity of caspase-3, -7, -8, and -9 was measured with luminescent caspase activity assays. Expression of nuclear factor-κB (NF-κB) gene was assessed with RT–quantitative (q)PCR. Cytochrome c release apoptosis assay kit was used to extract mitochondria and cytosol. Cytochrome c release, cleavage of Bid, and expression of B-cell lymphoma 2 (Bcl-2) were determined by Western blot.

Results:
Cell death ELISA revealed that ACh is able to reduce Fas-induced apoptosis in keratocytes. Analysis of the activity of effector caspases-3 and -7 showed that ACh, when added to Fas-treated cells, decreases the activation of both these enzymes. The activity of initiator caspases -8 and -9 also decreased when ACh was added to Fas-treated cells. This antiapoptotic effect of ACh was dependent on ACh concentration and activation of muscarinic ACh receptors. Analysis of the antiapoptotic mechanisms triggered by ACh showed that ACh downregulates expression of FasL-induced NF-κB RNA expression, upregulates expression of antiapoptotic protein Bcl-2, downregulates expression of proapoptotic protein Bad, reduces cytochrome c release, and prevents proapoptotic Bid protein cleavage.

Conclusions:
Acetylcholine has an antiapoptotic effect in a Fas-apoptosis model of human primary keratocytes in vitro. It is therefore possible that ACh may play a role in corneal wound healing, by modulating its initiation phase.

The cornea is the avascular highly organized and specialized transparent tissue covering the front part of the eye, protecting the eye from infectious agents and refracting light onto the retina.1 The stroma is the main component of the cornea, comprising its middle, thickest layer, and is an avascular tissue sparsely populated by cells called keratocytes and composed of mostly collagen I and V fibers arranged into bundles called lamellae.2 The keratocytes are quiescent, mesenchymal-derived cells3,4 that produce and secrete extracellular matrix (ECM) components, contributing to corneal transparency. Upon injury to the cornea, the keratocytes convert into cells with a repair phenotype, either promoting healthy healing or inducing formation of scar tissue that decreases corneal transparency, leading to corneal opacity and subsequent vision loss.5 Corneal blindness is the fourth leading cause of blindness globally, with ocular trauma and corneal ulcerations being its main etiology.6 Corneal opacities may arise from diseases such as keratitis, for example, by herpes simplex infection,7 trauma to the eye (e.g., chemical burn),8 or after various surgeries, like cataract surgery,9 and are a result of impaired healing of the cornea.

The healing of the corneal stroma occurs in different phases: keratocyte apoptosis, inflammatory phase, proliferation and migration phase, and ECM remodeling phase.10 In the initial phase, apoptosis of keratocytes adjacent to the wound leaves a zone that is free of cells, and it has been considered the initiator of the stromal healing, most probably controlling the entire process.11,12 Apoptosis is triggered by interleukin-1 (IL-1) released by injured epithelium.13 However, other studies have also implicated the Fas ligand system as a possible modulator of keratocyte apoptosis in response to injury.12 Thus, injured epithelium has been shown to produce Fas ligand in response to injury, which in turn could trigger apoptosis of keratocytes.14 Fas ligand (FasL, CD95L) is a type II transmembrane protein belonging to the tumor necrosis factor (TNF) superfamily.15 It initiates an extrinsic apoptotic pathway through trimerization of the death receptor Fas (CD95) on cell surfaces16 and leads to autoactivation of the enzyme caspase-8, which then triggers activation of other caspases such as caspase-3, -7, and -9.17 Caspases are a family of cysteine proteases, which exist as inactive proenzymes that become activated upon cleavage. They are divided into two classes: the initiator caspases (caspase-2, -8, -9, and -10) and the effector caspases (caspase-3, -6, and -7).18 Initiator caspases are at the beginning of the caspase signaling pathways and are responsible for activating the effector caspases, which execute cell death.19 The intrinsic apoptotic pathway is mediated by mitochondria and, in response to apoptotic stimuli,20 it leads to release of cytochrome c from the intermembrane space of the mitochondria into the cytoplasm, which subsequently activates caspase-9. Activation of the Fas pathway also leads to cytochrome c release through cleavage of proapoptotic mitochondrial protein Bid.21 Bid is a member of the B-cell lymphoma 2 (Bcl-2) family, which upon apoptotic stimuli is cleaved by caspase-8, and its carboxyl terminal p15 fragment then translocates to the outer membrane of the mitochondria and eventually triggers cytochrome c release.22 Moreover, nuclear factor-κB (NF-κB), a transcription factor, which is involved in signals typically leading to cell growth or cell differentiation,23 has been shown to play a crucial role in apoptosis regulation.24

Finding ways to reduce or even prevent keratocyte apoptosis has become an important issue. Regulation of caspase activation can provide a survival signal in order for the cells to withstand the apoptotic stimuli. One way of controlling caspase activity is through regulation of mitochondrial proteins involved in apoptosis. One such protein is the antiapoptotic Bcl-2. It belongs to the family of Bcl-2 proteins, which might exert either proapoptotic or antiapoptotic properties25; Bcl-2 inhibits release of cytochrome c from the mitochondria,26 and it has been implicated in modulation of mitochondrial calcium homeostasis and proton flux.27 Bad, a proapoptotic protein and a member of the Bcl-2 family, associates with Bcl-2.28 Phosphorylation of Bad at Ser112 and Ser136 inhibits its apoptotic activity through preventing its association with Bcl-2.29

Acetylcholine (ACh) is a classical neurotransmitter that via nicotinic or muscarinic acetylcholine receptors (nAChRs and mAChRs, respectively) mediates chemical neurotransmission in order to optimize an organism's mood and survival.30 However, ACh regulates a wide variety of cellular responses outside the neuronal system. Thus, ACh has been found to be involved in regulation of immune function of T cells,31 proliferation of embryonal carcinoma cells,32 and neuronal differentiation.33 Interestingly, stimulation of both mAChRs and nAChRs has been shown to protect many cell types from apoptosis,32,34,35 and moreover, it has been shown that ACh can prevent apoptosis of embryonic stem cells and cardiomyocytes through regulation of Bcl-2.36,37 Therefore, of relevance for apoptosis of keratocytes in the corneal stroma, and its role in corneal wound healing, is the fact that ACh and its receptors have been found in human keratocytes both in situ in corneal tissue sections and in vitro in primary human cell cultures,38 and that ACh also enhances proliferation of human keratocytes in vitro.39

In the present study, we hypothesize that ACh reduces Fas-induced apoptosis of keratocytes, through stimulation of either nAChRs or mAChRs and possibly via regulation of initiator caspases as well as regulation of mitochondrial proteins involved in apoptosis. By doing so, endogenous ACh might modulate corneal wound healing and prevent and/or reduce corneal scar formation.

Materials and Methods

Preparation of Human Corneas

Healthy human corneas were stored in a corneal biobank at the University Hospital of Umeå, Sweden. The corneas were obtained from deceased individuals who had chosen, when alive, to donate their corneas post mortem for transplantation or research, according to Swedish law. The corneas were delivered for research purposes if they were not used for transplantation. The Regional Ethical Review Board in Umeå reviewed the study and determined it to be exempt from the requirement for approval (2010-373-31M). The study was completed in accordance with the principles of the Declaration of Helsinki.

The isolation and culture of primary keratocytes was performed as previously described.38 Donated corneas were scraped with a scalpel in order to remove epithelial and endothelial cells. Next, the central part of the cornea was separated from the peripheral part, and both were minced with a scalpel. Samples were digested with 2 mg/mL collagenase diluted in DMEM/F-12 + GlutaMAX medium containing 2% FBS and 1% penicillin-streptomycin (DMEM/F-12 2% FBS), overnight at 37°C. The samples were then centrifuged and resuspended in DMEM/F-12 2% FBS, transferred to culture dishes, and cultured at 37°C with 5% CO2. Culture medium was changed every second to third day until cells reached confluence. Trypsin-EDTA (0.05%) was used to detach the confluent cells, and the cells were split in a 1:2 ratio. Central keratocytes in passages 4 to 5 were used in this study. DMEM/F-12 2% FBS was used to propagate the cell culture, whereas DMEM/F-12 0.1% FBS was used for both seeding of the cells and performing the experiments. The corneas were assessed individually.

Apoptosis Measurements

Cells (104 per well) were seeded in 96-well plate in DMEM/F-12 0.1% FBS and left to adhere for 24 hours at 37°C. Next, cells were treated with 10−5 M atropine for 30 minutes, 100 nM mecamylamine for 30 minutes, or left untreated. Afterward, cell death was induced with 250 ng/mL FasL, and ACh was added simultaneously to appropriate wells. Apoptosis was detected after 24 hours with cell death detection ELISA PLUS, according to manufacturer's instructions. Briefly, cells were lysed and placed into a streptavidin-coated 96-well plate. Anti-histone-biotin and anti-DNA-POD antibodies were added and samples were incubated. Amount of nucleosomes retained in the immunocomplexes was determined by measuring absorbance with a plate reader (BioTek, Winooski, VT, USA) at 405 nm (reference at 490 nm).

TUNEL Assay

Cells (0.5 × 106 per well) were seeded in triplicate in six-well plates in DMEM/F12 0.1% FBS and left to adhere for 24 hours at 37°C. Next, cell death was induced with 250 ng/mL Fas ligand, and 10−8 M ACh was added simultaneously to appropriate wells. DNA strand breaks induced in apoptotic cells were detected after 24 hours with APO-BrdU TUNEL Assay kit according to manufacturer's instructions. Briefly, cells were detached from the plate, centrifuged, and fixed in 1% paraformaldehyde for 15 minutes on ice. Next, fixed cells were washed with PBS, resuspended in 70% ice-cold ethanol, and stored in a −20°C freezer for 12 to 18 hours. Next, samples were washed, resuspended in DNA-labeling solution containing reaction buffer, TdT (terminal deoxynucleotidyl transferase) enzyme, BrdUTP (deoxythymidine analogue 5-bromo-2′-deoxyuridine 5′-triphosphate), and water and incubated for 60 minutes at 37°C in a water bath. Afterward, samples were washed and incubated with BrdU-Alexa Fluor 488 antibody for 30 minutes at room temperature. Samples were analyzed with LSR II flow cytometer (BD Biosciences, Franklin Lakes, NJ, USA) and FlowJo software (FlowJo, Ashland, OR, USA).

Caspase Activity

Cells (104 per well) were seeded in 96-well plates in DMEM/F-12 0.1% FBS and left to adhere for 24 hours at 37°C. Next, cells were treated with 10−5 M atropine for 30 minutes, 100 nM mecamylamine for 30 minutes, or left untreated. Afterward, cell death was induced with 250 ng/mL FasL, and ACh was added to appropriate wells. Cells were incubated for 1, 2, and 3 hours (caspase-8), 1, 2, 3, and 4 hours (caspase-9), or 1, 2, 3, and 8 hours (caspase-3/-7) at 37°C. Caspase-3/-7, caspase-8, and caspase-9 activity assays were used to determine caspase activities according to manufacturer's instructions. Briefly, caspase reagent was added to each sample and incubated for 1 hour at room temperature on a shaker. Luminescence was measured using a plate reader (BioTek), and caspase activities were determined.

Isolation of Mitochondrial and Cytosolic Fractions and Cytochrome Release

Cells (5 × 106) were plated in 10-cm Petri dishes in DMEM/F-12 supplemented with 0.1% FBS and left to adhere for 24 hours at 37°C. Cell death was induced with 250 ng/mL, and 10−8 M ACh was added to appropriate plates. Cells were incubated for 3 and 24 hours. Cytochrome c release apoptosis assay kit was used to extract mitochondria and cytosol according to manufacturer's instructions. Briefly, cells were collected and washed with ice-cold PBS. Cells were resuspended in cytosol extraction buffer containing dithiothreitol (DTT) and protease inhibitors. Next, cells were centrifuged at 10,000g for 30 minutes at 4°C, and cytosolic fraction was collected as supernatant. The remaining pellet was resuspended in mitochondrial extraction buffer containing DTT and protease inhibitors. Protein content was measured with Bradford assay (Bio-Rad, Hercules, CA, USA; 5000006). Five micrograms of each fraction was loaded, separated on 12% SDS-PAGE gel, and transferred to polyvinylidene fluoride (PVDF) membrane. Membranes were blocked with 5% nonfat dry milk or 5% BSA for 1 hour at room temperature and incubated with cytochrome c or Bid primary antibody at 4°C overnight. Next, the membranes were incubated with HRP-conjugated secondary antibody for 1 hour at room temperature. Images were taken by Odyssey Fc imaging system (LI-COR, Lincoln, NE, USA).

Western Blot

Cells (2.5 × 105/well) were plated in six-well plates in DMEM/F-12 0.1% FBS and left to adhere for 24 hours at 37°C. Cell death was induced with 250 ng/mL Fas, and ACh was added to appropriate wells. Cells were incubated for 3, 6, 12, and 24 hours. Next, cells were washed with PBS and freeze/thawed three times and lysed in radioimmunoprecipitation assay (RIPA) buffer supplemented with proteinase and phosphatase inhibitor cocktail (Thermo Fisher Scientific, 10085973). Protein concentration was assessed by Bradford assay (Bio-Rad). Samples were separated on SDS-polyacrylamide gels and transferred to PVDF membranes. Membranes were blocked with 5% BSA for 1 hour at room temperature and incubated with primary antibodies (β-actin, Bcl-2, Bad, and phospho-Bad) at 4°C overnight. Next, the membranes were incubated with HRP-conjugated secondary antibody for 1 hour at room temperature. Images were taken by Odyssey Fc imaging system (LI-COR). Densitometry was performed using ImageJ analysis software (http://imagej.nih.gov/ij/; provided in the public domain by the National Institutes of Health, Bethesda, MD, USA).

MTS Assay

Cells (0.3 × 104/well) were plated in 96-well plate in DMEM/F-12 0.1% FBS and left to adhere for 24 hours at 37°C. Next, 10−8 M ACh was added to appropriate wells, and cells were incubated for 6 days. Medium and ACh were exchanged every second day. On day 6, apoptosis was induced with 250 ng/mL FasL. On day 7, cell viability was measured with CellTiter 96 Aqueous One Solution Assay according to manufacturer's instructions. Briefly, 20 μL of the solution was added to the cells at day 7 and incubated for 2 hours at 37°C. Cell viability was measured at absorbance of 490 nm with a plate reader.

Reverse Transcription–Polymerase Chain Reaction (RT-qPCR)

Cells (2.5 × 105/well) were plated in six-well plates in DMEM/F-12 supplemented with 0.1% FBS and left to adhere for 24 hours at 37°C. Cell death was induced with 250 ng/mL, and 10−8 M ACh was added to appropriate wells. Cells were incubated for 2, 6, 12, and 24 hours. RNA extraction kit was used according to manufacturer's instructions in order to extract total RNA. RNA (400 ng) was reverse transcribed to cDNA with a high-capacity cDNA reverse transcription kit. Nuclear factor-κB mRNA levels were determined by using NFKB1 gene expression probe. Samples were run in duplicates in ViiA 7 Real-Time PCR System (Thermo Fisher Scientific); 18S probe served as an endogenous control (Thermo Fisher Scientific; 4333760F). Analysis was performed with ViiA 7 Software (Thermo Fisher Scientific).

Statistical Analysis

All experiments were performed in triplicate. Data are presented as mean ± SD. Statistical analysis was carried out using 1-way ANOVA and Bonferroni post hoc test. Differences were considered statistically significant at a P value of < 0.05. The experiments were performed successfully at least three times; that is, at least three separate experiments were performed with cells isolated from different patients.

The potential antiapoptotic effect of ACh in Fas-induced apoptosis was assessed by Cell Death ELISA PLUS and APO-BrdU TUNEL assay. The results showed that keratocytes undergo apoptosis when treated with FasL. Fas-induced apoptosis was decreased when ACh was added to the cells together with FasL. The antiapoptotic effect was dependent on ACh concentration. Acetylcholine at 10−8 and 10−7 M significantly reduced Fas-induced apoptosis; 10−6 M ACh had no antiapoptotic effect (Fig. 1A). The antiapoptotic effect of ACh was further confirmed by TUNEL assay. The results showed that ACh decreased the number of Fas-induced BrdU-Alexa Fluor 488–positive cells when ACh was added to the cells together with FasL (Fig. 1B). In order to assess if mAChRs and/or nAChRs are involved in the antiapoptotic effect of ACh, keratocytes were pretreated with the mAChR antagonist atropine or with the nAChR antagonist mecamylamine. Atropine at 10−5 M abolished the antiapoptotic ACh effect (Fig. 1C). In contrast, 100 nM mecamylamine did not affect the antiapoptotic effect of ACh. Interestingly, 10−6 M ACh showed an antiapoptotic effect after blockage of nAChRs (Fig. 1D), something not seen when treated with 10−6 M ACh alone.

Our recently published study showed that ACh enhances keratocyte proliferation.39 In order to study whether ACh-enhanced proliferation may lead to enhanced keratocyte transdifferentiation into apoptosis-resistant cells, keratocytes were treated with 10−8 M ACh for 6 days. On day 6, apoptosis was induced with 250 ng/mL FasL. The results showed that keratocytes treated with ACh undergo apoptosis, and this process seemed to be more pronounced than in untreated cells (Supplementary Fig. S1). Moreover, prolonged treatment of keratocytes with ACh did not alter expression of keratocan (keratocyte marker) or expression of α-SMA (α-smooth muscle actin, marker of transdifferentiated myofibroblasts) (data not shown).

In order to assess whether ACh can decrease effector caspase-3 and caspase-7 activity after Fas-induced apoptosis, specific caspase-3/-7 activity assay was used. Caspase-3/-7 activities were assessed 1, 2, 3, and 8 hours after apoptosis induction. Fas ligand increased activity of caspase-3 and caspase-7 as early as 2 hours. This activity was decreased significantly when 10−8 M ACh (2, 3, and 8 hours after apoptosis induction) and 10−7 M ACh (8 hours after apoptosis induction) were added to the cells together with FasL. However, adding 10−6 M ACh did not cause decrease of caspase-3/-7 activity (Fig. 2A). Additionally, cells were pretreated with 10−5 M atropine or 100 nM mecamylamine to examine if mAChR and/or nAChR activation are necessary for the ACh inhibiting effect on Fas-induced capsase-3 and caspase-7 activity at 8 hours after apoptosis induction. The ACh effect was no longer present when mAChRs were blocked (Fig. 2B). However, caspase-3 and caspase-7 activity was decreased by ACh even after blocking nAChRs, indicating that the antiapoptotic effect of ACh was not blocked by mecamylamine (Fig. 2C).

Caspase-8 is known to initiate an apoptotic pathway triggered by external stimuli. To test whether ACh is able to regulate caspase-8 activity and therefore control the process of apoptosis, specific caspase-8 activity was analyzed. Caspase-8 activity was assessed 1, 2, and 3 hours after apoptosis induction. Fas ligand increased activity of caspase-8 after 3 hours. This activity was decreased significantly when 10−8 and 10−7 M ACh was added to the cells together with FasL. Acetylcholine at 10−6 M had no effect on caspase-8 activity (Fig. 3A). To examine if mAChR and/or nAChR activation are necessary for ACh-induced caspase-8 decrease, cells were treated with 10−5 M atropine and/or 100 nM mecamylamine. Caspase-8 activity was determined 3 hours after apoptosis induction. The ACh effect on caspase-8 activity was no longer present when mAChRs were blocked, resulting in higher activity of caspase-8 (Fig. 2B). Blocking nAChRs still resulted in decreased caspase-8 activity when 10−8 M ACh was used. However, 10−7 M ACh showed no effect on caspase-8 activity after mecamylamine treatment, meaning that nAChRs might play a minor role in ACh-induced regulation of caspase-8 (Fig. 3C).

ACh decreases initiator caspase-8 activity through mAChRs after Fas-induced apoptosis. (A) Apoptosis was induced in 104 keratocytes/well with 250 ng/mL FasL. 10−8, 10−7, or 10−6 M ACh was added simultaneously to Fas-treated cells. Cells were incubated for 1, 2, or 3 hours at 37°C. Caspase-8 activity was measured with a luminescent caspase assay. ACh decreased caspase-8 activity in a dose-dependent manner. (B) 104 keratocytes/well were pretreated with 10−5 M atropine for 30 minutes at 37°C. Next, 250 ng/mL FasL was added followed by addition of 10−8, 10−7, or 10−6 M ACh. Cells were incubated for 3 hours at 37°C. Caspase-8 activity was measured with a luminescent caspase assay. Treatment of cells with atropine and ACh did not result in caspase-8 activity decrease. (C) 104 keratocytes per well were pretreated with 100 nM mecamylamine (Mec) for 30 minutes at 37°C. Next, 250 ng/mL FasL was added followed by addition of 10−8, 10−7, or 10−6 M ACh. Cells were incubated for 3 hours at 37°C. Caspase-8 activity was measured with a luminescent caspase assay. Treatment of cells with mecamylamine and 10−8 M ACh resulted in caspase-8 activity decrease. Treatment of cells with mecamylamine and 10−7 or 10−6 M ACh did not result in caspase-8 activity decrease. Values are means ± SD. n.s., not significant; *P < 0.05; **P < 0.01.

Figure 3

ACh decreases initiator caspase-8 activity through mAChRs after Fas-induced apoptosis. (A) Apoptosis was induced in 104 keratocytes/well with 250 ng/mL FasL. 10−8, 10−7, or 10−6 M ACh was added simultaneously to Fas-treated cells. Cells were incubated for 1, 2, or 3 hours at 37°C. Caspase-8 activity was measured with a luminescent caspase assay. ACh decreased caspase-8 activity in a dose-dependent manner. (B) 104 keratocytes/well were pretreated with 10−5 M atropine for 30 minutes at 37°C. Next, 250 ng/mL FasL was added followed by addition of 10−8, 10−7, or 10−6 M ACh. Cells were incubated for 3 hours at 37°C. Caspase-8 activity was measured with a luminescent caspase assay. Treatment of cells with atropine and ACh did not result in caspase-8 activity decrease. (C) 104 keratocytes per well were pretreated with 100 nM mecamylamine (Mec) for 30 minutes at 37°C. Next, 250 ng/mL FasL was added followed by addition of 10−8, 10−7, or 10−6 M ACh. Cells were incubated for 3 hours at 37°C. Caspase-8 activity was measured with a luminescent caspase assay. Treatment of cells with mecamylamine and 10−8 M ACh resulted in caspase-8 activity decrease. Treatment of cells with mecamylamine and 10−7 or 10−6 M ACh did not result in caspase-8 activity decrease. Values are means ± SD. n.s., not significant; *P < 0.05; **P < 0.01.

Caspase-9 is an initiator caspase that can be activated either by extrinsic (via caspase-8) or by intrinsic stimulus. To examine whether ACh is able to regulate caspase-9 activity, specific caspase-9 activity was analyzed. Caspase-9 activity was analyzed 1, 2, 3, and 4 hours after induction of apoptosis. Fas ligand increased activity of caspase-9 at 3 and 4 hours. This activity was decreased significantly when 10−8 and 10−7 M ACh were added to the cells together with FasL. Acetylcholine at 10−6 M had no effect on caspase-9 activity (Fig. 4A). Involvement of mAChRs and nAChRs in this process was determined 4 hours after apoptosis induction. The ACh inhibitory effect on Fas-induced caspase-9 activity was still present after mAChRs were blocked when 10−8 M ACh was used. This was not the case for 10−7 M ACh, for which blocking of mAChRs with 10−5 M atropine abolished the ACh effect (Fig. 4B). Similarly, blocking nAChRs with 100 nM mechamylamine resulted in abolished ACh-induced inhibition of Fas-induced caspase-9 activity when 10−7 M ACh was used, but not when 10−8 M ACh was used (Fig. 4C).

Induction of NF-κB has been shown to have both anti- and proapoptotic effects. To test whether FasL or ACh is able to induce expression of NF-κB, RT-qPCR was performed at 2, 6, 12, and 24 hours after induction of apoptosis (Fig. 5A). The results show that FasL significantly increased expression of NF-κB at 6, 12, and 24 hours. However, 10−8 M ACh treatment alone did not show any change in the gene expression. Fas ligand–induced NF-κB expression was significantly decreased when 10−8 M ACh was added to the cells together with FasL. Bcl-2 exerts an antiapoptotic function through inhibition of mitochondrial cytochrome c release and therefore caspase-9 activation. To test whether the ACh antiapoptotic effect is driven by regulation of this protein, Western blot analysis was performed (Fig. 5B). Keratocytes were treated with FasL and/or ACh (10−8 or 10−7 M) for 3, 6, 12, and 24 hours. The results show that expression of Bcl-2 was decreased after induction of apoptosis with FasL as compared to control at all time points tested. Bcl-2 expression was significantly increased when 10−8 M (at 6, 12, and 24 hours) and 10−7 M (at 3, 12, and 24 hours) ACh were added to the cells together with FasL as compared to FasL alone. Again, ACh alone upregulated expression of Bcl-2 as compared to control (10−8 M at 3, 6, 12, and 24 hours, and 10−7 M at 6, 12, and 24 hours) (Supplementary Fig. S2A). To further explore the mechanism behind ACh-driven Bcl-2 increased expression, expression of Bad, a proapoptotic protein that inactivates Bcl-2, and expression of its phosphorylated form, which has antiapoptotic properties, were assessed by Western blot (Fig. 5C). Keratocytes were treated with FasL and/or ACh (10−8 or 10−7 M) for 3, 6, 12, and 24 hours. The results showed that expression of Bad is increased 3 and 6 hours after induction of apoptosis with FasL as compared to control. Bad expression was significantly decreased when 10−8 and 10−7 M ACh were added to the cells together with FasL as compared to FasL alone at 3 and 6 hours after apoptosis induction (Supplementary Fig. S2B). Moreover, the results showed that phosphorylation of Bad, which prevents association of Bad with Bcl-2, is decreased after induction of apoptosis with FasL as compared to control at all time points tested. Phospho-Bad expression was significantly increased when 10−8 M (at 3, 6, 12, and 24 hours) and 10−7 M (at 3, 6, and 12 hours) ACh were added to the cells together with FasL as compared to FasL alone (Supplementary Fig. S2C). Cytochrome c, under normal conditions, is associated with the inner membrane of mitochondria. Upon apoptotic stimuli it is released to the cytoplasm where it binds to apoptotic protease-activating factor 1 (Apaf-1) and activates caspase-9 and downstream caspases. In order to assess whether decreased caspase-9 activation after FasL/ACh treatment of keratocytes is caused by decreased cytochrome c release, cellular fractionation was performed. Keratocytes were treated with FasL and/or 10−8 M ACh for 3 hours. Mitochondrial and cytosolic fractions were isolated and subjected to cytochrome c Western blot. The results show that FasL treatment resulted in release of cytochrome c from mitochondria to the cytoplasm. However, when ACh was added to the cells together with FasL, the cytochrome c release to the cytoplasm was decreased. Moreover, cleavage of the proapoptotic protein Bid was assessed. It could be observed that FasL treatment induced cleavage of Bid, whereas when ACh was added to the cells together with FasL, cleavage of Bid occurred to a lesser extent at both 3 and 24 hours of treatment (Fig. 5D).

This study shows that ACh reduces Fas-induced apoptosis in human keratocytes, and that this antiapoptotic effect is mostly mediated through activation of mAChRs. The Fas apoptosis model has been chosen due to the fact that, although it is known that FasL is not expressed in keratocytes of unwounded cornea,40 when the cornea gets injured, IL-1 released from the corneal epithelium stimulates keratocytes to express FasL, which in turn triggers apoptosis in keratocytes that express the Fas receptor.14 This ACh effect is driven by its ability to downregulate the FasL-induced NF-κB gene expression. Moreover, ACh regulates activities of caspases through reduction of Bid protein cleavage, phosphorylation of the proapoptotic Bad protein, and upregulation of caspase-9 inhibitory protein Bcl-2, which ultimately leads to decreased cytochrome c release from mitochondria. All in all, this mechanism leads to diminished keratocyte death.

Previous reports have demonstrated that activation of either mAChRs or nAChRs leads to protection against apoptosis. Thus, activation of nAChRs has been shown to protect against apoptosis of neurons through activation of protein kinase B,41 and to inactivate proapoptotic mitochondrial protein BAD in small cell lung cancer cells.42 Muscarinic AChR activation has been reported to protect human neuroblastoma cells from apoptosis triggered through DNA damage and oxidative stress35 and to block caspase activation in rat phaochromocytoma cells (PC12 cells).34 Moreover, it has been shown that M1, M2, and M3 subtypes of mAChRs trigger the antiapoptotic pathway through activation of protein kinase B.43 Additionally, activation of M1 and M3 subtypes results in inhibition of caspase-2 and caspase-3 in Chinese hamster ovary cells, PC12 cells, and NG108-15 (hybrid of mouse neuroblastoma and rat glioma cells).34,44,45 The mAChR-mediated antiapoptotic effect is also dependent on transcription of the antiapoptotic protein Bcl-2, which can be induced by mAChR subtype M3.46 In the present study, we have observed that in human keratocytes, ACh was able to decrease Fas-induced cell death and that this process was dependent on activation of mAChRs. We think that the reason behind muscarinic receptors being responsible for transducing the ACh antiapoptotic signal is the fact that the muscarinic receptors are abundantly expressed by keratocytes. In our previous study,38 we showed that cultured keratocytes express M1, M3, M4, and M5 subtypes of muscarinic receptors (lacking only M2). We also observed that the M3 receptor was most probably responsible for transducing the ACh-enhanced proliferation signal.39 There are no data available on expression of nicotinic receptors in keratocytes, but it is known that nicotinic receptors are expressed abundantly in corneal epithelium.47 In our lab we have studied expression of the α7 nicotinic receptor and found that it was not expressed in keratocytes (data not published).

The higher concentration of ACh used (10−6 M) did not have an antiapoptotic effect. An equivalent phenomenon was observed in our earlier studies on ACh-enhanced proliferation of keratocytes, in which only lower concentrations of ACh (10−8 and 10−7 M) enhanced keratocyte proliferation, whereas 10−6 M ACh did not.39,48 This might be due to the fact that higher concentrations of ACh cause downregulation of mAChRs leading to decreased sensitivity of the receptors.49,50 As reported before,39,48 we have observed downregulation of mAChR subtype M3 after treatment of keratocytes with a high concentration of ACh (10−6 M). Therefore, as subtype M3 has been implicated in an antiapoptotic effect, this could explain why we do not observe such an effect when the higher concentration of ACh is used. Moreover, we have shown that the ACh-enhanced keratocyte proliferation does not lead to apoptosis-resistant keratocytes and that the prolonged treatment of keratocytes with ACh does not alter their phenotype.

Our results in the present study furthermore showed that ACh decreases effector caspase-3 and caspase-7 activities after Fas-induced cell death, which results in decreased apoptosis. Caspases are intracellular proteases that usually propagate apoptosis, proliferation, inflammation, migration, and differentiation.51 They are produced by the cell and kept in their inactive forms until receipt of specific stimuli, such as FasL.52 The caspases are divided into initiator caspases and effector caspases. The initiator caspases (caspase-2, -8, -9, and -10) cleave inactive forms of effector caspases (caspase-3, -6, and -7). The effector caspases act directly on specific cellular substrates to trigger the apoptotic process.53 Our results showed that ACh is able to decrease activities of initiator caspase-8 and caspase-9. It has been reported that increasing endogenous ACh levels decrease caspase 8 activity in rats,54 and activation of nAChRs has been shown to inhibit caspase-9 in mouse astrocytes.55 Our results suggest that the ACh-induced decrease of caspase-8 activity is mediated primarily through activation of mAChRs, as blocking of these receptors abolished the antiapoptotic effect of all ACh concentrations tested. However, it seems that nAChRs might also be involved in parts of this process, as the antiapoptotic effect of ACh on caspase-9 seemed to be partially abolished by the nAChR blocker at one ACh concentration (10−7 M). This study further shows that the decrease of caspase-9 activity is mediated through both the mAChRs and nAChRs. As this caspase is part of both the intrinsic and extrinsic apoptotic pathways, the activating stimuli might come from different sources. Other studies have also shown that regulation of caspase-9 activity can be mediated by nAChRs55 or mAChRs.46 All in all, ACh seems to act as a pan-caspase inhibitor; that is, it has an inhibitory effect on both initiator and effector caspases.

It has been reported that induction of apoptosis in mouse in vivo cornea models results in an immediate keratocyte death, but in vitro response to apoptotic stimuli occurs after several hours.13 This phenomenon could also be observed in our study. We think that the differences between the in vitro and in vivo apoptotic time course might occur due to several factors related to isolation of keratocytes and their further culturing. This issue has been also addressed in several studies.13,56 Factors such as plate support (plastic, coated with collagen, silicone) also affect keratocyte phenotype and physiology. In our lab we have observed that keratocytes grown on different supports exhibit different expression of markers and receptors. Experimental setup might also play a role, as cells grown in vitro will be of different density than cells in the cornea, and this might affect their response to the stimuli. Cells in vivo are in a specific, complex environment, surrounded by other cells and ECM; their faster response to stimuli might also be due to combined “forces” of many substances present, and it is necessary for the in vivo system to work fast in order to, for example, prevent spreading of an infection.

Nuclear factor-κB, a transcription factor, which is generally regarded as antiapoptotic, has been recently shown to promote this process,57,58 especially when cellular stress, such as serum withdrawal or ultraviolet radiation, is applied.59 Our results showed that FasL induces increased NF-κB gene expression. Therefore, the results suggest that NF-κB has proapoptotic properties in this model of cell death of keratocytes. Our results further show that ACh is able to reduce the upregulation of the proapoptotic NF-κB, resulting in partial protection from apoptosis. Bcl-2, the antiapoptotic member of the evolutionarily conserved Bcl-2 family, is central to the control of cell survival, as it is able to directly affect mitochondrial events leading to the activation of caspases.60 Specifically, it inhibits cytochrome c release from mitochondria and subsequently inhibits caspase-9 activation26; therefore it can be considered as a caspase-9 regulator. It has been reported that activation of mAChRs leads to upregulation of Bcl-2 in neural cells61 and embryonic stem cells.36 Our results show that ACh is able to control expression of Bcl-2 protein and decrease the release of cytochrome c from mitochondria, thereby controlling activation of caspase-9 to further increase keratocyte survival after induction of apoptosis. Bcl-2 expression could be observed both in apoptosis-induced cells to which ACh was added and in cells treated with ACh alone. Additionally, it has previously been shown that Bcl-2 inhibits NF-κB activity by controlling its nuclear expression in human embryonic kidney cell line 293.62 Therefore, one could speculate that ACh-increased Bcl-2 expression might downregulate the FasL-induced NF-κB gene expression in this apoptosis model. The enhanced expression of Bcl-2 could be explained by the fact that ACh leads to phosphorylation of the proapoptotic protein Bad, leading to its inability to associate with Bcl-2 and to transduce the apoptotic signal. Lastly, we show that ACh reduces cleavage of the proapoptotic protein Bid. As Bid cleavage results in cytochrome c release and subsequent caspase-9 activation,63 this property of ACh further enhances its antiapoptotic effect.

The cornea is normally well protected from injury and infection. However, due to trauma or disease it may become injured. The normal healing process will lead to a healthy, transparent cornea, but any disruption or dysregulation in this process might cause scar formation and subsequent vision impairment.64 The initial phase of the wound healing process, which is the apoptosis of keratocytes adjacent to the wound, is considered the initiator of stromal healing.11,12 Our results suggest that ACh is able to significantly decrease the apoptosis of keratocytes. Hence, it is possible that ACh might be involved in the control of this very important step and in regulating the normal corneal wound healing, possibly promoting a healthy and transparent cornea, as interfering with the onset of the wound healing process would prevent the subsequent activation of keratocytes and decrease the possibility of scar formation. However, in the case of corneal infection, keratocyte apoptosis is beneficial as it prevents the spread of the infection. Therefore, decreasing it or abolishing it with ACh would not be desirable. We think that ACh could possibly be used for preventing unwanted keratocyte death during corneal surgeries. However, the role of ACh on myofibroblasts (fibrosis) has not been studied; hence, its application or suitability in patients with already developed fibrosis should be further investigated.

In conclusion, ACh triggers an antiapoptotic mechanism in a Fas-induced apoptosis model for human keratocytes. This mechanism is activated mostly through mAChRs. Moreover, the results indicate that ACh regulates apoptosis through downregulation of FasL-induced NF-κB gene expression and through control of initiator caspase-8 and -9 activity. The antiapoptotic effect is further exerted through inhibition of Bid cleavage and upregulation of Bcl-2. These events lead to decreased cytochrome c release and caspase-9 activity, ultimately resulting in decreased activity of effector caspase-3 and -7 and therefore in less cell death. Control of keratocyte apoptosis during injury to the cornea is of great importance to proper wound healing. Therefore, ACh might be involved in the control of corneal wound healing, which, properly regulated, leads to decreased formation of scars and thus less subsequent loss of vision.

Acknowledgments

The authors thank Jialin Chen, Sandrine Le Roux, and Gabor Borbely for technical and scientific advice. We would also like to thank Maria Brohlin, Randi Elstad, and Berit Byström for help in providing the donated corneas from the biobank.

Supported by the national Swedish Research Council (Grant no. 521-2013-2612), the J.C. Kempe and Seth M. Kempe Memorial Foundations (JCK-1222), the Swedish Society of Medicine (SLS-410021/-504541), the Cronqvist Foundation (SLS-249071/-329561/-596181), the foundation Kronprinsessan Margaretas Arbetsnämnd för synskadade KMA (2012/26, 2013/10), and Västerbotten County Council “Spjutspetsmedel” (VLL-363161), and via federal funds through a regional agreement between Umeå University and Västerbotten County Council (VLL-549761).

ACh decreases initiator caspase-8 activity through mAChRs after Fas-induced apoptosis. (A) Apoptosis was induced in 104 keratocytes/well with 250 ng/mL FasL. 10−8, 10−7, or 10−6 M ACh was added simultaneously to Fas-treated cells. Cells were incubated for 1, 2, or 3 hours at 37°C. Caspase-8 activity was measured with a luminescent caspase assay. ACh decreased caspase-8 activity in a dose-dependent manner. (B) 104 keratocytes/well were pretreated with 10−5 M atropine for 30 minutes at 37°C. Next, 250 ng/mL FasL was added followed by addition of 10−8, 10−7, or 10−6 M ACh. Cells were incubated for 3 hours at 37°C. Caspase-8 activity was measured with a luminescent caspase assay. Treatment of cells with atropine and ACh did not result in caspase-8 activity decrease. (C) 104 keratocytes per well were pretreated with 100 nM mecamylamine (Mec) for 30 minutes at 37°C. Next, 250 ng/mL FasL was added followed by addition of 10−8, 10−7, or 10−6 M ACh. Cells were incubated for 3 hours at 37°C. Caspase-8 activity was measured with a luminescent caspase assay. Treatment of cells with mecamylamine and 10−8 M ACh resulted in caspase-8 activity decrease. Treatment of cells with mecamylamine and 10−7 or 10−6 M ACh did not result in caspase-8 activity decrease. Values are means ± SD. n.s., not significant; *P < 0.05; **P < 0.01.

Figure 3

ACh decreases initiator caspase-8 activity through mAChRs after Fas-induced apoptosis. (A) Apoptosis was induced in 104 keratocytes/well with 250 ng/mL FasL. 10−8, 10−7, or 10−6 M ACh was added simultaneously to Fas-treated cells. Cells were incubated for 1, 2, or 3 hours at 37°C. Caspase-8 activity was measured with a luminescent caspase assay. ACh decreased caspase-8 activity in a dose-dependent manner. (B) 104 keratocytes/well were pretreated with 10−5 M atropine for 30 minutes at 37°C. Next, 250 ng/mL FasL was added followed by addition of 10−8, 10−7, or 10−6 M ACh. Cells were incubated for 3 hours at 37°C. Caspase-8 activity was measured with a luminescent caspase assay. Treatment of cells with atropine and ACh did not result in caspase-8 activity decrease. (C) 104 keratocytes per well were pretreated with 100 nM mecamylamine (Mec) for 30 minutes at 37°C. Next, 250 ng/mL FasL was added followed by addition of 10−8, 10−7, or 10−6 M ACh. Cells were incubated for 3 hours at 37°C. Caspase-8 activity was measured with a luminescent caspase assay. Treatment of cells with mecamylamine and 10−8 M ACh resulted in caspase-8 activity decrease. Treatment of cells with mecamylamine and 10−7 or 10−6 M ACh did not result in caspase-8 activity decrease. Values are means ± SD. n.s., not significant; *P < 0.05; **P < 0.01.